The H+-ATPase HA1 of Medicago truncatula Is Essential for Phosphate Transport and Plant Growth during Arbuscular Mycorrhizal Symbiosis

H+-ATPase HA1 of Medicago truncatula Is Essential for Phosphate Transport and Plant Growth during Arbuscular Mycorrhizal Symbiosis. A key feature of arbuscular mycorrhizal symbiosis is improved phosphorus nutrition of the host plant via the mycorrhizal pathway, i.e., the fungal uptake of Pi from the soil and its release from arbuscules within root cells. Ef ﬁ cient transport of Pi from the fungus to plant cells is thought to require a proton gradient across the periarbuscular membrane (PAM) that separates fungal arbuscules from the host cell cytoplasm. Previous studies showed that the H + -ATPase gene HA1 is expressed speci ﬁ cally in arbuscule-containing root cells of Medicago truncatula . We isolated a ha1-2 mutant of M. truncatula and found it to be impaired in the development of arbuscules but not in root colonization by Rhizophagus irregularis hyphae. Arti ﬁ cial microRNA silencing of HA1 recapitulated this phenotype, resulting in small and truncated arbuscules. Unlike the wild type, the ha1-2 mutant failed to show a positive growth response to mycorrhizal colonization under Pi-limiting conditions. Uptake experiments con ﬁ rmed that ha1-2 mutants are unable to take up phosphate via the mycorrhizal pathway. Increased pH in the apoplast of abnormal arbuscule-containing cells of the ha1-2 mutant compared with the wild type suggests that HA1 is crucial for building a proton gradient across the PAM and therefore is indispensible for the transfer of Pi from the fungus to the plant.


INTRODUCTION
Arbuscular mycorrhizal (AM) symbiosis between obligate biotrophic fungi of the phylum Glomeromycota and the majority of land plants is based upon bidirectional nutrient transfer between host plants and AM fungi (Smith and Smith, 2012). AM symbiosis is ancient and is believed to have facilitated colonization of land by aquatic plants ;450 million years ago (Redecker et al., 2000). The widespread occurrence of AM symbioses today (Smith and Read, 2008) indicates that they continue to play key roles in terrestrial ecology. Growth and spore development of AM fungi depends on successful colonization of roots to access plant carbohydrates and convert them into fatty acids and other compounds (Solaiman et al., 1999;Trépanier et al., 2005). In return, AM fungi take up minerals, especially Pi, from the soil and deliver them to their host plants (Marschner and Dell, 1994). Pi availability in the soil is known to regulate AM fungal colonization of the root, and in low-P soils, AM fungi can provide most of the P needed by plants (Bucher, 2007;Smith et al., 2011;Yang and Paszkowski, 2011). AM fungi can also transfer soil N and S to plants under some conditions (Leigh et al., 2009;Casieri et al., 2012;Koegel et al., 2013;Sieh et al., 2013).
Exchange of phosphate and carbohydrates between AM fungi and colonized plant cells is mediated by specific transporters on the fungal membrane and the surrounding plant periarbuscular membrane (PAM) (Smith and Smith, 1990). Specific plant phosphate transporters (PTs) are confined to the PAM, indicating that arbuscules are the main site of Pi transfer to the plant (Harrison et al., 2002). By contrast, localization of a fungal monosaccharide transporter to arbuscules and intraradical hyphae indicates that fungal uptake of plant sugar may not be restricted to arbuscules (Helber et al., 2011). Downregulation of either the plant phosphate transporter or the fungal monosaccharide transporter resulted in reductions not only of arbuscule abundance, but also of overall hyphal spread in the cortex of the root (Javot et al., 2007;Helber et al., 2011).
The PAM-located PT is a phosphate/H + cotransporter. H + -ATPase activity has been observed at the PAM (Marx et al., 1982), which presumably generates the proton gradient required for phosphate transport from the periarbuscular space into the plant cytoplasm via the PAM-PT. An H + -ATPase gene, HA1, has been identified in Medicago truncatula that is expressed specifically in arbuscule-containing cells of mycorrhizal roots (Krajinski et al., 2002). To test the hypothesis that HA1 is required to energize phosphate transport to the plant, we isolated an ha1-2 mutant of M. truncatula and compared its symbiotic phosphate uptake and growth characteristics to those of the wild-type.

Identification of an Insertion Mutant for HA1
An ha1-2 mutant was identified among 8000 long terminal repeat retrotransposon Tnt1-insertion lines of M. truncatula, via a PCRbased screen (see Methods). Sequencing of the mutant allele revealed a Tnt1 insertion in exon 8 of HA1 (Figure 1). Homozygous ha1-2 progeny were isolated from a self-pollinated heterozygous ha1-2/HA1-2 individual. Homozygous HA1-2 progeny of the same parent were also isolated and used in subsequent experiments as wild-type controls. The effect of the Tnt1 insertion on HA1 transcription was determined by RT-PCR. Using HA1-specific primers flanking the Tnt1 insertion site, a 1123-bp cDNA fragment was amplified from wild-type plants ( Figure 1). Surprisingly, a shorter amplicon was produced from cDNA of homozygous ha1-2 mutant lines ( Figure 1). Subsequent sequencing of the PCR product from the ha1-2 mutant revealed the complete absence of exon 8. Therefore, insertion of Tnt1 resulted in splicing out of exon 8 in the ha1-2 mutant. Exon 8 consists of 207 bp encoding 69 amino acids of transmembrane domains 3 and 4 that are essential for H + -ATPase activity of related proteins (Palmgren, 2001;Morth et al., 2011) (Supplemental Figure 1). Therefore, the ha1-2 mutant can be regarded as a complete loss-of-function mutant. As expected due to the restricted expression of HA1 in mycorrhizal roots, nonmycorrhizal ha1-2 mutants did not show any phenotypes.
ha1-2 Mutants Are Colonized by R. irregularis but Do Not Show a Positive Growth Response and Exhibit Altered Arbuscule Morphology Four weeks after inoculation with R. irregularis, more than 90% the root system of wild-type plants were colonized by the AM fungus ( Figure 2). The roots of ha1-2 mutant plants did not show significant differences with regard to colonization frequency and hyphae and arbuscule numbers ( Figure 2). Fungal colonization resulted in a large Roots of mycorrhizal wild-type plants exhibited typical arbuscules with fine hyphal branches, under both low-and high-Pi conditions ( Figure 3). By contrast, only truncated arbuscules were observed in roots of the ha1-2 mutant under both conditions ( Figure 3). Consistent with containing smaller, truncated arbuscules, levels of R. irregularis rRNA were 20-fold lower in roots of the mutant than of the wild-type (fungal RNA in Supplemental Figure 2).

Decreased Expression of Mycorrhiza-Inducible Genes in Colonized ha1-2 Mutant Roots
Expression of eight plant genes known to be induced in mycorrhizal roots and accumulation of R. irregularis rRNA were analyzed in roots of the ha1-2 mutant. Transcripts for all of these plant genes and fungal rRNA were significantly less abundant in the ha1-2 mutant than in the wild type; this included transcripts of PT4 (Javot et al., 2007) and other genes identified previously to be highly induced in mycorrhizal roots (Wulf et al., 2003;Pumplin et al., 2010;Hogekamp et al., 2011;Gaude et al., 2012b;Devers et al., 2013;Supplemental Figure 2).
Artificial MicroRNA-Mediated HA1 Silencing also Leads to Truncated Arbuscules To confirm that the aberrant arbuscule morphology in ha1-2 mutant plants was due to loss of HA1 function, we employed the artificial microRNA (amiR) technique to reduce HA1 expression in M. truncatula roots. Expression of HA1-amiR was driven by the arbuscule-specific Mt-PT4 promoter (Harrison et al., 2002), and (A) Mycorrhizal colonization parameters of R. irregularis colonized roots of ha1-2 and wild-type plants grown low Pi condition. Plants were harvested 4 weeks after inoculation and mycorrhizal colonization parameters (Trouvelot et al., 1986) were estimated. Data represent mean of two (wild type) and four (ha1-2) replicates 6 SD. Significant differences between the wild type and ha1-2 were not detected. (B) Shoot fresh weights of ha1-2 and wild-type plants. Mycorrhizal plants (myc) were inoculated with R. irregularis. All plants were harvested 3 weeks after inoculation (nm, nonmycorrhizal plants). Shown are mean values and standard deviations of three biological replicates. Different letters indicate fresh weight values that differ significantly from each other (P < 0.05). transgenic roots were identified by coexpression of the fluorescent protein DsRED. Following fungal inoculation, control roots expressing DsRED alone produced fully developed, highly branched arbuscules (Supplemental Figure 3). By contrast, no such arbuscules were detected in roots transformed with the HA1-amiR construct and in which HA1 transcript levels were <20% of control levels. R. irregularis rRNA accumulation and Mt-PT4 transcript levels were also significantly decreased in HA1-amiR roots. However, transcript levels of two other H + -ATPase genes expressed in M. truncatula roots (Krajinski et al., 2002) were not affected in the HA1-amiR roots.

HA1 Is Not Essential for the Root Nodule Symbiosis
Analysis of nodulated roots and nodules of ha1-2 mutant and wild-type plants revealed no induction of HA1 transcript during root nodule symbiosis (Supplemental Figure 4).
To investigate if HA1 is essential for efficient root nodule symbiosis, we inoculated ha1-2 and wild-type plants with Sinorhizobium meliloti. Four weeks after inoculation, wild-type and ha1-2 plants showed increased shoot fresh weight as a consequence of symbiotic nitrogen fixation and established root nodule symbiosis (Supplemental Figure 5). The two plant lines showed (E) and (F) Arbuscule phenotype of the wild type and ha1-2, respectively, at higher magnification. Fifteen optical sections were superimposed to represent complete arbuscules. a, arbuscules; ih, intercellular hyphae. Bars = 40 µm. (G) Percentages of wild-type or ha1-2 root fragments showing mature and degenerated arbuscules. Root fragments were scored for the presence of fully developed, mature arbuscules, and degenerated arbuscules. Root fragments containing both types of arbuscules were counted for both categories. Data represent mean and 6 SD of three biological replicates with at least 15 root fragments counted for each plant. Different letters indicate significantly different values (P < 0.01).
ha1-2 Mutants of M. truncatula similar nodule numbers and levels of Mt-N24 (Godiard et al., 2007) expression. These data indicate that HA1 is not necessary for efficient root nodule symbiosis.

HA1 Is Required for the Uptake of Phosphate via the Mycorrhizal Pathway
To test whether HA1 is required for the uptake of phosphate via the mycorrhizal pathway, we set up compartmented systems wherein the plant and the hyphal compartment were separated by nylon meshes and an air gap. Half of the plants were inoculated with R. irregularis. Twelve weeks after inoculation, extraradicular fungal hyphae had passed the air gap and colonized the hyphal compartment. At this time point, 33 PO 4 32 was added to the hyphal compartment and plants were harvested at 15 weeks after inoculation. No differences in the frequency of mycorrhizal colonization were observed between wild-type plants and ha1-2 mutants (62.5 6 3.1 and 59.5 6 5.6, respectively). Moreover, hyphal length density (HLD) was not significantly different between mutants and wild-type plants; the HLD of R. irregularis was 4.8 cm g 21 and 4.2 cm g 21 soil dry weight when associated with the wild-type and mutant plants, respectively. Nonmycorrhizal control plants did not show a significant uptake of 33 P. By contrast, 33 P accumulated in both shoots and roots of mycorrhizal wild-type plants, indicative of 33 P uptake via the mycorrhizal pathway. Roots and shoots of mycorrhizal ha1-2 mutants did not show significant uptake of 33 P (Figure 4), confirming that mycorrhizal ha1-2 mutants were unable to take up phosphate via a mycorrhizal pathway. Therefore, we can assume that HA1 is essential for mycorrhizal phosphate uptake in M. truncatula.

HA1 Is Required for Production of an Acidic Compartment in Arbuscule-Containing Cells
HA1 is the only M. truncatula H + -ATPase-encoding gene that is strongly induced in arbuscule-containing cells. Strong expression of an H + -ATPase-encoding gene in arbuscule-containing cells presumably results in an electrochemical proton gradient and thus in acidification of apoplastic compartments. Therefore, a functional knockout should result in reduced acidification of apoplastic spaces in arbuscule-containing cells. To test this, we applied the pH-sensitive Lysosensor DND-189 dye to stain mycorrhizal roots of wild-type and ha1-2 mutants ( Figure 5). This acidotropic probe accumulates in acidic compartments, where protonation at pH 5.2 leads to a green fluorescence. Staining of mycorrhizal wild-type roots revealed strong fluorescence in the arbuscule-containing cells of the inner cortical cell layer ( Figure 5A). Subsequent application of carbonylcyanid-m-chlorophenylhydrazon (CCCP), which acts as protonophor, led to a strong reduction of the fluorescence confirming that the observed fluorescence after Lysosensor DND-189 staining results from acidification in the corresponding compartments ( Figure 5B). By contrast, fluorescence was barely detectable in mycorrhizal ha1-2 mutant roots, pointing to diminished acidification around the abnormal arbuscules ( Figure 5C). Laser scanning microscopy confirmed that arbuscule-containing cells of wild-type roots showed strong fluorescence, whereas arbusculecontaining cells of ha1-2 mutant roots did not ( Figures 5D and 5E). This indicates HA1 is required for the production of an acidic compartment in arbuscule-containing cells. ha1-2 mutants and wild-type plants were inoculated with R. irregularis and grown in compartmented systems. 33 P was added to the hyphal compartment 12 weeks after inoculation, and plants were harvested 15 weeks after inoculation. The data shown are average values of four independent replicates; error bars represent standard deviations. Different letters indicate statistical differences.

DISCUSSION
Here, we have shown that the H + -ATPase HA1 of M. truncatula, which is strongly induced in arbuscule-containing cells, is required for arbuscule development. Disruption of the gene in ha1-2 mutants leads to truncated arbuscules and decreased acidification of apoplastic spaces in arbuscule-containing cells. The ha1-2 mutants also show strongly impaired uptake of phosphate by the mycorrhizal uptake pathway. From this, we conclude that HA1 is essential for P transport via the mycorrhizal uptake pathway.
Phosphorus is often the main limiting factor for plant growth. One major advantage of the AM symbiosis for plants is the more effective uptake of phosphate via the mycorrhizal pathway (Smith et al., 2011), which often results in increased growth of mycorrhizal plants under low-P conditions. AM symbiosis increased the growth of wild-type M. truncatula plants by more than threefold (shoots) under low-P conditions, whereas AM symbiosis had no effect on the growth of ha1-2 mutant plants. Uptake experiments in compartmented systems showed that ha1-2 mutants were unable to take up phosphate via the mycorrhizal pathway. The uptake of phosphate from the periarbuscular space presumably requires a proton gradient generated by HA1. The acidic nature of the periarbuscular space in functional arbuscule-containing cells has been shown previously (Guttenberger, 2000). We found that arbuscule-containing cells of ha1-2 mutants showed an increased pH in the apoplast as a consequence of lacking HA1 proton pumping activity. Therefore, it can be concluded that HA1 is essential for phosphorus transfer from AM fungi to the plant. A model explaining the role of HA1 in phosphate uptake into plants is shown in Figure 6.
Reduced phosphate transport across the PAM as a result of reduced H + -pumping across this membrane in the ha1-2 mutant probably also explains the abnormal arbuscule development observed in the mutant (Figure 3). Truncated arbuscules were similarly observed in rice mutants defective for Os-HA1, which also encodes a H + -ATPase induced during AM symbiosis (Wang et al., 2014). Moreover, two other M. truncatula ha1 mutant alleles have been identified (Wang et al., 2014), confirming that the phenotype described herein results from a mutation in the HA1 gene. Mutation in PT4 of M. truncatula, which encodes a phosphate transporter located in the PAM, also resulted in a phenotype with degenerated arbuscules (Javot et al., 2007). Moreover, a similar phenotype was described in rice (Oryza sativa) defective for the Pt11 phosphate transporter (Yang et al., 2012). Thus, there appears to be a phosphate-sensitive checkpoint in the host plant cell that aborts the resource-intensive arbuscule development (Gaude et al., 2012a) at an early stage if phosphate is not transferred from the fungus to the plant.
Similar arbuscule morphology was reported recently for stunted arbuscule (str) mutants, which are defective in two half-ABC Phosphate is transferred from the fungal cytoplasm across the fungal membrane in arbuscule-containing cells by a yet unidentified transport mechanism and released to the periarbuscular space. Specific PT proteins (blue) are located in the periarbuscular membrane and mediate the transport of phosphate across this membrane. Plant PTs are phosphate/proton symporters. Thus, a proton gradient is essential for phosphate uptake into arbuscule-containing cells. In M. truncatula, this proton gradient is generated by a H + -ATPase protein (HA1; orange). ha1-2 Mutants of M. truncatula transporters (Zhang et al., 2010;Gutjahr et al., 2012). However, str mutants showed drastically reduced fungal colonization of roots, in contrast to the ha1-2 mutant in which the degree of colonization was normal (Figure 2). The reduced fungal colonization of str mutants might be explained by lack of transport of unknown signal molecule(s), the substrate(s) of the ABC transporters (Zhang et al., 2010). If this is the case, then HA1 is not required for such signaling. HA1 is clearly also not required for nutrient supply to the invading fungus, which shows normal hyphal growth in the roots of the ha1-2 mutant (Figures 2 and 3). This is in contrast to M. truncatula pt4 mutants, in which fungal growth is severely impaired (Javot et al., 2007). It is assumed that functional PT4 proteins are required to signal the presence of a beneficial AM fungus and to allow fungal growth (Javot et al. 2007). R. irregularis-colonized ha1-2 mutants show significantly reduced PT4 expression, which, however, might be sufficient to sustain these signaling events and thus sustain fungal growth.
The normal level of fungal colonization in ha1-2 mutant roots indicates that colonization is not dependent on mature arbuscules, which are absent in the mutant. This assumption is supported by the finding that plants defective for a vesicle-associated membrane protein required for PAM synthesis show truncated arbuscules with very few branches but normal levels of intraradical colonization (Ivanov et al., 2012). AM fungi receive much of the carbon from their plant host in the form of sugars, and at least one fungal hexose transporter has been found on the intraradical fungal hyphae in addition to the arbuscule (Helber et al., 2011). Here, our results with the ha1-2 mutant indicate that carbon transport across the PAM in mature arbuscule-containing cells is not required for hyphal growth. No differences in the frequency of mycorrhizal colonization were observed between wild-type plants and ha1-2 mutants, and hyphal length density was also not significantly different between the mutants and wild-type plants. Since the majority of fungal carbon is derived from the host plants, our observations suggest that carbon transport from the plant to the fungus is not dependent on prior phosphate transport from the AM fungus. However, carbon transport across the PAM in mature arbuscule-containing cells might still be important for fungal metabolism within the arbuscule.
The assumption that fungal growth and especially arbuscule development are not strictly dependent on phosphate transfer to the host plant is also supported by pt4 mutants, which show normal arbuscules when grown under nitrogen-replete conditions (Javot et al., 2011). In addition, strong asymmetry in carbon investment by the plant and the phosphate provided by the AM fungus has been observed in common mycorrhizal networks (Walder et al., 2012). This and the phenotype observed here of unaltered fungal growth and impaired mycorrhizal phosphate uptake challenges the prevailing view that the plant regulates fungal colonization according to the amount of phosphate it receives from the fungus.

Isolation of an ha1-2 Mutant
To identify a Medicago truncatula mutant carrying a Tnt1 insertion in HA1, we screened ;8000 plants with Tnt1 insertions (Pislariu et al., 2012). Using two pairs of nested primers (Supplemental Table 1), we identified a M. truncatula mutant plant with a Tnt1 insertion in exon 8 of HA1.

Plant Growth and Inoculation with Rhizophagus irregularis
M. truncatula seed germination was performed as described (Branscheid et al., 2010). Seedlings were transplanted into pots containing a mixture of expanded clay and silica sand. For inoculation with R. irregularis, an inoculum was mixed with the growth substrate (1:10 v/v). The inoculum was obtained by growing Allium schoenoprasum with R. irregularis as described previously (Mrosk et al., 2009). Unless otherwise indicated, all plants were grown in a greenhouse at 24°C with a 16-h-light/8-h-dark cycle. Plants were fertilized with half-strength Hoagland solution (Hoagland and Arnon, 1950) containing either 20 µM or 1 mM phosphate, twice per week.

DNA and RNA Extraction
RNA of roots and leaves was extracted using the Invisorb Spin Plant Mini kit (Invitek). Genomic DNA was isolated using the DNeasy Plant Mini kit (Qiagen).

Quantitative RT-PCR and RT-PCR
Quantitative RT-PCR was performed as described recently (Branscheid et al., 2010). Oligonucleotide sequences of all primers are listed in Supplemental  Table 1. The amplification efficiencies (E) were calculated using the LinRegPCR program (Ramakers et al., 2003). At least three biological replicates were performed for each quantitative RT-PCR analysis.

Staining of Fungal Structures
Roots were stained with wheat germ agglutinin coupled to AlexaFluor488 according to Gaude et al. (2012b). Images were collected on a Leica TCS-SP5 confocal microscope (Leica Microsystems) using a 633 water immersion objective with a numerical aperture of 1.2, zoom 1.6. AlexaFluor488 was excited at 488 nm, and emitted light was collected from 505 to 582 nm. Optical sections were acquired at 0.3-to 0.5-µm intervals. Images were processed using ImageJ software (Wayne Rasband, National Institutes of Health).

Phosphate Uptake Experiments
The phosphate uptake experiments were set up using compartmented microcosms (Koegel et al., 2013), wherein one plant and one hyphal compartment were connected, but separated by two 21-mm nylon meshes and an air gap in between. The air gap was created by placing two 5-mm plastic meshes between the two 21-mm nylon meshes. The two compartments were filled with sterile sand (quartz sand from Alsace, 0.125 to 0.25 mm; Kaltenhouse) and zeolithe (2:1 w/w). M. truncatula seedlings (the wild type and ha1-2 mutants) were inoculated with a 2-g (;100 spores) inoculum of R. irregularis BEG-75 or with 2 g of sterilized (120°C, 20 min) inoculum as a nonmycorrhizal control. In the center of the hyphal compartment, a 21-mm nylon mesh bag of 15 mL was inserted and kept empty until introduction of the 33 P-labeled substrate 12 weeks after inoculation. Then, the nylon mesh bag was filled with 13 g sand labeled with 750 kBq 33 PO 4 32 (Hartmann Analytic). Two milliliters of water was added to wet the 33 P-spiked sand without inducing mass flow. The microcosms were irrigated with distilled water twice a week. In addition, the compartments were amended weekly with 8 mL of Long Ashton nutrient solution. Plants were grown under controlled conditions (16 h light at 28°C and 8 h dark at 15°C, constant relative aerial humidity of 65%). Plants were harvested 15 weeks after inoculation. The root colonization was estimated by a modified line intersection method (McGonigle et al., 2006). Shoot and root samples were dried for 24 h at 105°C and weighed separately. Dried shoots and roots were ground at 30 Hz in a mixer mill (MM2224; Retsch). Phosphorus was extracted by acid digestion (Murphy and Riley, 1962), and 33 P contents were measured using a Packard 2000 liquid scintillation counter (Hewlett-Packard). Mean comparisons were performed by independent paired t tests for 33 P uptake and for root colonization.
HLD HLD was measured separately for the root and hyphal compartments and was determined by the modified grid-line intersection method (Jakobsen et al., 1992) using 10 g of the growth substrate. After sieving successively through a 400-and a 32-µm mesh, the material was collected and transferred into 50 mL of distilled water and homogenized for 10 s at full speed in a blender. The suspension was transferred into a beaker, diluted to 500 mL, and stirred for 1 min before five subsamples were taken every 10 s and loaded onto the Filtration apparatus (MF-Membrane filter 1.2 µm; Millipore).

Staining of Acidic Compartments
Roots of mycorrhizal wild-type and ha1-2 plants were sectioned into 200-to 300-µm-thick sections by hand. Sections were transferred into a solution consisting of 3 µM Lysosensor DND-189 (Invitrogen) and 15 µg/mL wheat germ agglutinin conjugated with tetramethylrhodamine for localization of fungal structures in 10 mM MOPS/KOH buffer (pH 8.0) and incubated in the dark at room temperature overnight. As a control, stained root sections were incubated with 50 µM CCCP in 10 mM MOPS/KOH buffer for 20 min. Sections were analyzed either by epifluorescence microscopy using a multi-zoom microscope AZ-100 (Nikon) equipped with the proper filter cube (EX460-500/ 505/BA510) or by confocal microscopy using an LSM700 (Zeiss). For the latter, the excitation wavelengths 488 and 555 nm were used, and fluorescence light was collected in the range of 493 to 544 nm and 560 to 630 nm for Lysosensor DND-189 and WGA-tetramethylrhodamine, respectively. All settings (laser power, pinhole diameter, and detector gain) remained constant for all pictures.

amiR Silencing
An HA1-amiR sequence (59-39: TTGTTAAGTAACATGAAGCCC) was designed using the WMD3 microRNA designer tool (Ossowski et al., 2008), inserted into the mtr-miR159b backbone by an overlapping PCR strategy including parts of the pBluescript II SK cloning site and subcloned into pCR2.1-TOPO (Invitrogen). For amiR expression in M. truncatula roots, a binary vector (pRed-Pt4exp, accession number JX280943) that includes the DsRED expression cassette of pRedRoot (Limpens et al., 2004) as visible marker for root transformation was used . AmiR expression was driven by the Mt-PT4 promoter (Harrison et al., 2002) that enables strong expression in arbuscule-containing cells. The resulting pRed-PT4 pro :amiR-HA1 vector was transformed into M. truncatula roots using Agrobacterium rhizogenesmediated transformation as described recently (Gaude et al., 2012b).

Statistics
To test for differences between plant genotypes and treatments, data were analyzed by ANOVA followed by Tukey HSD test or by Student's t test for pairwise comparisons using the Sigmaplot software package (Systat).

Accession Numbers
Sequence data from this article can be found in the GenBank/EMBL databases under accession number AJ132891.1 (HA1).

Supplemental Data
The following materials are available in the online version of this article.